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Targeted Large-scale Deletion of Bacterial Genomes Using CRISPR-Nickases Kylie S Standage-Beier, Qi Zhang, and Xiao Wang ACS Synth. Biol., Just Accepted Manuscript • DOI: 10.1021/acssynbio.5b00132 • Publication Date (Web): 09 Oct 2015 Downloaded from http://pubs.acs.org on October 10, 2015

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ACS Synthetic Biology

Targeted Large-scale Deletion of Bacterial Genomes Using CRISPR-Nickases Kylie Standage-Beier1, 2, Qi Zhang2, Xiao Wang2* 1

School of Life Sciences, 2School of Biological and Health Systems Engineering, Arizona State University, Tempe, AZ 85287, USA. Abstract Programmable CRISPR-Cas systems have augmented our ability to produce precise genome manipulations. Here we demonstrate and characterize the ability of CRISPR-Cas derived nickases to direct targeted-recombination of both small and large genomic regions flanked by repetitive elements in E.coli. While CRISPR directed double-stranded DNA breaks are highly lethal in many bacteria, we show that CRISPR-guided nickase systems can be programmed to make precise, nonlethal, single-stranded incisions in targeted genomic regions. This induces recombination events and leads to targeted deletion. We demonstrate that dualtargeted nicking enables deletion of 36 and 97 Kb of the genome. Furthermore, multiplex targeting enables deletion of 133 Kb, accounting for approximately 3% of the entire E.coli genome. This technology provides a framework for methods to manipulate bacterial genomes using CRISPR-nickase systems. We envision this system working synergistically with preexisting bacterial genome engineering methods.

Keywords: CRISPR, Cas9 nickase, genome engineering, chromosome deletion, direct repeats, recombination Large chromosomal rearrangements and deletions have been observed in both natural and laboratory bacterial evolution studies1–3 and shown to have profound impacts on bacterial physiology, such as improved bioproduct production4, increased strain fitness5, or changed tolerance to stress6. However, such desired genotypes can only be produced in the lab by time-consuming and laborious directed evolution experiments. This is because bacterial genome rearrangement and deletion events only occur when a spontaneous but stochastic DNA break emerges between direct-repeat sequences3. Efficient methods to target this process for large-scale genome manipulations would be beneficial to biotechnology. Current genome rearrangement strategies often rely on the insertion of exogenous recombinase sites into the bacterial genome to facilitate remodeling7–12. The use of site-specific recombinases provide efficient tools for in vivo DNA manipulation, however, targeting recombination in a programmable and controllable fashion would broaden and simplify implementations of genome engineering for more applications. Recent advances in using the Clustered Regularly Interspaced Short Palindromic Repeats (CRISPR) and CRISPR associated (Cas) systems for precise genome editing13–18 have paved the way of using it for large scale genome modifications. CRISPR functions as a prokaryotic and archaeal immune system protecting cells from foreign nucleic acids19–21. CRISPR RNAs (crRNAs) direct degradation of target DNAs through simple Watson-

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Crick base pairing to recruit Cas9 nuclease to introduce double stranded DNA breaks (DSBs) to foreign nucleic acids20–22. These systems have minimal sequence targeting constraints, requiring only a small proto-spacer adjacent motif (PAM) sequence23,24, which makes it possible to program CRISPR to target almost any sequence. CRISPR induced chromosomal DSBs are highly lethal to many bacteria15,25–28 and leads to rapid target DNA degradation26. This has been exploited to select for desired mutations15,29 and also enabled genome engineering methods in bacteria that are capable of DSB repair30,31. However, current genomic DNA manipulations in E.coli largely employ site-specific recombinases, which can facilitate recombination between defined recombinase sites. These methods have enabled large-scale manipulations of the bacterial genome including deletion7,8, inversion7, and insertion10,32 of DNA. Employing CRISPR-Cas systems to target recombination between arbitrary sequences would complement these approaches to increase the versatility of rationally designed genome manipulations. Type II CRISPR systems containing Streptococcus pyogenes Cas9 nuclease have been shown to enable user designated single-stranded DNA cleavage (nicking). This is accomplished through mutation of RuvCI or HNH nuclease domains33. Methods employing nicking versions of Cas9 have been utilized in eukaryotic organisms to direct homologous recombination (HR) and reduce off-target mutations13,34,35. As genomic DSBs are highly lethal to many bacterial species, we sought to investigate the potential application of nicking CRISPR systems to induce recombination on the bacterial chromosome. Here, we demonstrate that nicking versions of Cas9 are non-lethal when targeting the wild type bacterial genome. Short guide RNAs (sgRNAs)33 can be programmed to direct single-stranded DNA breaks (SSBs) resulting in host cell recombination. We functionally validate this with nicking mutant Cas9D10A using a fluorescent reporter system. We show that dual-targeting sgRNAs to genomic repeat sequences leads to recombination across 36 to 97 Kilobases. The easily implementable and modular nature of our of CRISPR plasmid system enables multiplex targeting. Multi-targeted CRISPR-nickase systems can remodel the E.coli genome resulting in 133 Kb of deleted DNA, leading to changes of multiple phenotypes. To experimentally assess the lethality of CRISPR-directed SSBs in wild type E.coli, we constructed a two-plasmid system (Supplemental Fig. 1): one plasmid expresses Cas9 and another expresses an sgRNA against a genome integrated green fluorescent protein (GFP) gene (Fig. 1A). Multiple mutant versions of Cas9, including Cas9D10A, Cas9H840A, and a Cas9D10A, H840A double mutant36, were tested. The D10A mutation abrogates function of the RuvCI nuclease domain, generating a Cas9 capable of only nicking the DNA strand complementary to the sgRNA. On the other hand, the H840A mutation removes function of the HNH nuclease domain resulting in a Cas9 that cleaves only the non-complementary strand to the sgRNA (Fig. 1a)33. Combination of both D10A and H840A leaves Cas9 catalytically dead (dCas9)36. Therefore, dCas9 can only bind to target DNA without any cleavage of the chromosome. We transformed GFP targeting sgRNA, sg(C1), into E. coli K12 coexpressing the


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aforementioned Cas9 variants and compared resulting colonies to a control non-targeting sgRNA, sg(-). As can be seen in Fig. 1b, sgRNA targeting GFP showed no viable colony forming units when wildtype Cas9 (Cas9WT) was expressed, while nicking or catalytically inactive variants of Cas9 don’t affect viability. This suggests that CRISPR induced DSBs are lethal to E. coli cells while CRISPR induced SSBs have no severe impact on viability. It is also demonstrated that such impacts are not cleavage position dependent (Supplementary Fig. 2). Although it has been reported that HR could be an inconvenience in building synthetic gene circuits with repetitive parts37,38, a precise nick between repeat sequences could hypothetically induce rationally designed HR to achieve targeted genome deletion39. To explore the potential of using CRISPR generated nicking to induce targeted HR, we constructed a genome integrated synthetic dual-reporter system (Fig. 2a) to query gene deletion and viability. In this system, GFP was designed to be flanked by two 129 base pair direct repeats (R). We systematically targeted nicking sites between direct repeats to both strands of GFP with Cas9D10A and Cas9H840A. The fluorescent population distributions of sgRNA(T1) Cas9D10A indicate that GFP is phenotypically lost in 86% of the cells (Fig. 2b, red). This target site is ~110bp away from the left repeat. It is interesting to see that the percentage of GFP negative cells quickly decreases as the target site is moved further away from the left repeat (Fig. 2c, blue triangles). However, Cas9H840A nicks result in less recombination (Fig. 2c). This suggests a strong strand and position dependency of single-nick induced local-homologous recombination. To verify HR as the cause of GFP signal loss, individual colonies were obtained from transformations for further analysis. Fluorescence microscope images show that, when targeted by sgRNA(T1) and Cas9D10A, cells have normal morphology and red fluorescent protein (RFP) expression while GFP fluorescence is undetectable (Fig. 2d, bottom), (for microscopy controls see Supplemental Fig. 3). This ruled out cell death as the cause of diminished GFP signal. Gel-electrophoresis results also verified a GFP knockout due to HR (Fig. 2e). Sanger sequencing of the PCR products further confirmed that recombination indeed occurred between the two direct repeats, yielding the same GFP deletion product (Supplemental Fig. 4). Finally, without the presence of two direct repeats on each end of GFP, neither Cas9D10A nor Cas9H840A can induce targeted GFP fluorescence abolishment (Supplementary Fig. 5). Furthermore, DSBs targeted between homologous sequences do not effectively lead to target deletion (Supplemental Fig. 6). Taken together, our results show that chromosomally targeted Cas9D10A can induce targeted SSBs and enable recombination between homologous sequences. To investigate how CRISPR-directed SSBs can be applied to delete large sections of genome in E.coli, we designed sgRNAs to target insertion sequences (IS) in direct repeat orientations and tested for genome deletion. Widespread distributions of transposable DNAs, such as IS elements, have been implicated to be responsible for large genomic changes1,40, and also provide a rich source of repetitive sequences to apply our method. We identified a region with two 1.2 Kb IS5 elements 35 Kb apart. The DNA flanked by


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IS5 elements contains 33 genes in total, including the Histidine (His) biosynthesis operon (Fig. 3a). The His operon provides us a convenient screenable marker for deletions. Cells containing the His operon are histidine prototrophs. When deleted, E.coli are converted to histidine auxotrophs (unable to grow without provision of the amino acid). To employ single-stranded lesions to direct recombination across the Histidine biosynthetic region, we systematically targeted a series of 10 sgRNAs (sgA-J) in the vicinity of the two IS5 sequences. These included targets directly adjacent to the repeats (R) on both strands along with multiple target sites within the 36.8 Kb region (Fig. 3a, b). We transformed K12 E.coli expressing nicking Cas9D10A with plasmids expressing individual and paired sgRNA combinations and cultured in pooled transformations and determined paired nicking did not lead to a substantial reduction in viability (Supplemental Fig. 7). We employed a PCR based approach to monitor recombination between repeats. Interestingly, the recombined site was undetectable in single targeting versions, unlike what’s shown above for GFP deletion. This indicates individual singlestrand breaks are insufficient to direct HR across large regions (Fig. 3c). Interestingly, when employing paired sgRNA, formation of the deletion product was detectable for multiple combinations (Fig. 3c). We isolated individual colonies from pooled transformations and subjected them to random screening for histidine auxotrophy. We determined sgRNAs targeting adjacent to each other, sg(A:B), and across the repetitive sequences, sg(B:I), had prominently undergone recombination (Fig. 3d). Our results show that sgRNAs generating a 5’ overhang across the repeat sequence, sg(B:I), and sgRNAs targeting in close proximity to each other , sg(A:B), have roughly 17±1.6% and 11±3.2% (±standard devation, 3 biological replicates) success rate of 36 Kb deletion, respectively (Fig. 3b d). Other paired-nicking transformations and sgRNA pair H:G were screened and confirm histidine operon deletions were rare (Supplemental Fig. 8 and 9). Finally, we confirmed that histidine auxotrophs consistently harbor the expected deletion by using multiple primer sets querying the presence of starting and recombined junctions (Fig. 3e). These results show that dual-targeted nicking systems can effectively induce recombination across 36 Kb with efficiencies approaching 20 percent of a bacterial population. The high efficacy of sg(B:I) provided us with design principles for further implementation of nicking-directed recombination systems. sgRNAs should be targeted adjacent to both ends of a repetitive sequence with Cas9D10A cut sites within close proximity (approximately 22 nucleotides) of the homology. The left sgRNA targets the + strand and the right guide targets the - DNA strand. This results in a 5’ overhang cleavage orientation (See Fig. 3b, 5’ OH). Interestingly, the requisite for a 5’ cut orientation is consistent with previously reported application of double nicking systems in mammalian cells34. To further explore the potential of our method to direct multiplexed large-scale genomic recombination, we identified a 97-kilobase genomic region with the tryptophan biosynthesis operon (TRP) and the pyrF gene flanked by identical 1.2 Kb IS5 repeats (Fig. 4a). This region enables screening for deletion by testing for tryptophan auxotrophy along with selection of deletion using 5-Fluoroorotic Acid (5-FOA)41. We created strain


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KSB1 with duplicate copies of essential genes found within the 97 Kb section of genome integrated into separate loci. This alleviated the essentiality of the region and enabled nickase-directed remodeling. To multiplex target the 97 Kb genomic region along with the 36 Kb region tested, we applied principles of dual sgRNA design from above experiments. sgRNAs K and L were designed and coupled to the previously employed guides B and I. This generated a fourtarget CRISPR system, sg(B:I:K:L), which should direct multiplex remodeling of the E.coli genome (Fig. 4a). We validated the necessity of cooperative nicking to induce homologous recombination and determined that four-targeted nicking did not substantially reduce transformation efficiency (Supplemental Fig. 10b, c). In pooled transformations of the multi-targeted CRISPR device, we detected recombination at both His and Trp regions (Fig. 4b). This indicated that HR at both sites was simultaneously occurring within the population of cells. To quantify the 97 Kb deletion we combined results for 5-FOA and TRP markers. 1.4±0.8% of cells exhibit 5-FOA resistance where 47% of the 5-FOA resistant colonies exhibited clear tryptophan auxotrophy (Supplementary Fig. 10d). Combining results for both markers, we estimated deletion efficiency to be 7 ± 4 deletions per 1000 cells. After initial 5-FOA/TRP screens we further tested for histidine deletion. Using stepwise screening (See Methods for details) we reliably obtained 36, 97 Kb single and 133 Kb dual-deletions from sg(B:I:K:L) transformations (Fig. 4c), indicating successful deletion of 133 Kb of E. coli genome using our approach. These results verify the dual-targeted sgRNA design for directed recombination. Likewise, the results exemplify the potential of multiplex CRISPR-guided nicking systems to remove large genomic regions. Here we demonstrate the ability for CRISPR targeted nicking systems to direct recombination across both small and large genomic regions. Furthermore, we systematically characterized various sgRNA target locations, choice of nicking Cas9, and overhang orientations to identify the optimal combination for efficient large-scale genome deletion. Targeting paired nicking across repeat elements in a 5’ overhang orientation with nick sites located within 40 bp of the homology sequence results in directed recombination. This enables a 3% reduction of the E. coli genome with only one plasmid transformation. Although the genome reductions reported here utilize repeat sequences to define the domain of remodeling, it is feasible to generate deletions of arbitrary position or to target genome integration by incorporation of plasmid based homologous sequences30,31. Genome rearrangements are an important part of bacterial evolution, which can facilitate large-scale organismal changes beneficial to biotechnology4–6. Using engineered SSBs, we enable recombination in bacterial cells resulting in multiplex genome rearrangements reaching 133 Kb in total, about 3% of the E. coli genome. In E.coli, genome reductions of such scale have been generated through multiple years of evolution1 and can be engineered through recombineering based methods12,42–44. Our system can recapitulate this scale of reduction through a single plasmid transformation. More importantly, the removal of these genome segments is targeted and its execution is controlled. It is conceivable that this method could assist in rationally designed and multi-targeted


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removal of pathways competing with the production of industrially useful chemicals and factors deleterious to growth. Collectively, our method provides a new tool for design and engineering of bacterial genomes45,46. Our results also provide a framework for methods that can change the landscape of bacterial genomes. Currently, genome-remodeling strategies in many bacteria often utilize site-specific recombinases7,47. These methods can direct high-efficiency recombination between recombinase sites. The current efficiency of CRISPR-nickase targeted deletion is relatively low, especially compared to efficiencies of site-specific recombinases, however these concerns may be addressed with improved sgRNA combinations and expression vectors. Nicking CRISPR systems, however, can be targeted with high versatility and leverage HR, which can utilize arbitrary sequences as substrates. Nicking CRISPR systems are also desirable to synthetic biology applications48–55. For instance, systems employing retron mediated reverse transcription have enabled genome directed rewriting and memory storage56, coupling CRISPRnickase systems to the output of inducible promoters could enable remodeling of the genome in response to environmental inputs. In summary, we present a strategy to overcome the cytotoxicity of genome targeting DSBs to make a platform for accurate large-scale bacterial genome manipulations. RNAprogrammed SSBs enable recombination resulting in bacterial genome deletion. With the growing availability of full genome sequencing, it is foreseeable that labs could identify an organism’s repetitive sequences in silico to predictably and combinatorialy direct large-scale remodeling to test strain characteristics for a myriad of basic science and engineering applications. Further extension of this method would enable researchers to target genome integration of synthetic gene pathways57, likewise, we envision this method working synergistically with preexisting genome engineering technologies. Methods: Strains and Media. Molecular cloning was conducted using E.coli NEB-10-Beta (New England Biolabs, NEB). Maintenance and cloning of vectors containing an R6Kγ origin of replication was conducted in a Pir+ E.coli strain . Fluorescent reporter experiments were conducted in K12 MG1655 (∆LacI ∆AraC) HK022(J04450 and/or I13522) (Registry of Standard Biological Parts). Initial 36 kilobase deletion experiments were conducted in K12 MG1655 ∆(LacI) ∆(AraC). Experiments involving the 97 kilobase deletion were conducted in strain “KSB1”, which is K12 MG1655 (American Type Culture Collection, ATCC, #700926) HK022(topA,cysB,ribA,fabI), 186(lapAB). Integrations into HK022 attB and 186 attB sites are described in strain construction section. LB Miller Medium (Sigma Aldrich, Sigma) was supplemented with appropriate antibiotics for plasmid maintenance: Ampicillin (100µg/ml) and/or Kanamycin (30µg/ml). Experiments involving fluorescent microscopy visualization of genome-integrated reporters used M9 Minimal Medium containing 0.2% glycerol. Screening for histidine and tryptophan auxotrophs was done on M9 Minimal Medium 0.4% glucose, 1.5% agar, 2 mg/ml Uracil with synthetic complete (SC) (ie. Leucine deficient supplement (Clonetech, #630414) + 200 mg/L Leucine (Sigma, #L8000), without histidine (-H) or without tryptophan (-T) amino acid supplement (Clonetech, #630415, #630414 respectively). 5-Fluoroorotic Acid


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(5-FOA, Gold Biotechnology, #F-230-5) media was made by supplementing 5-FOA to a final concentration of 600 mg/L and boiling to dissolve in LB agar miller medium. All cell culture was conducted at 37°C in either a stationary incubator for Petri dishes, or 250 RPM shaker for liquid media unless noted otherwise. Strain Construction. Genome integration of reporters and other genes were conducted using methods described previously32. Plasmids pOSIP-KH and pOSIP-KO were used to integrate into HK022 and 186 attB sites respectively. Fluorescent reporters (BioBrick #) parts RFP (J04450) and GFP (I13522) were cloned into the EcoRI and PstI sites of pOSIP-KH. The strain used for 97 Kb remodeling experiments was derived from ATCC, #700926. Integration vectors containing essential genes43,58 were created by stepwise cloning of topA:cysB, ribA, and fabI into pOSIP-KH using primers 18 through 23 with corresponding restriction sites indicated in Supplemental Table 2 and lapAB in pOSIPKO. The lapAB cassette was ligated and integrated in one step. Purified PCR products of primers 24 and 25 were digest with BamHI and SpeI and ligated into pOSIP-KO. Vectors were integrated by transforming ≤10 ng of plasmid into destination strains. Integrations were confirmed at HK022 att with primers 7 and 8, and 186 att with 26 and 27. R6Kγ vectors were propagated as plasmids at 30˚C and integrated at 37˚C. Construction of Cas9 Mutations. pCas9WT was obtained from Addgene (plasmid #44520)36. pCas9WT expresses S. pyogenes wild type Cas9 from a pLTetO promoter. The plasmid contains a high copy ColE1 origin of replication (~100 copies), the TetR repressor protein and an ampicillin resistance marker (Supplemental Fig. 1a). Cas9 D10A and H840A mutations were generated by PCR amplifying 10 pg of pCas9WT using primer pairs 3,4 and 5,6 respectively (Supplemental Table 2). PCR reactions were purified using GenElute PCR Cleanup Kit (Sigma, #NA1020) following the manufacturer’s protocol. DNA was digested with SapI and self-ligated with

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